A fluorogenic cyclic peptide for imaging and quantification of drug-induced apoptosis

A

fluorogenic cyclic peptide for imaging and

quanti

fication of drug-induced apoptosis

Nicole D. Barth

1,7

, Ramon Subiros-Funosas

1,7

, Lorena Mendive-Tapia

1

, Rodger Duf

fin

1

, Mario A. Shields

2

,

Jennifer A. Cartwright

1

, Sónia Troeira Henriques

3,6

, Jesus Sot

4

, Felix M. Goñi

4

, Rodolfo Lavilla

5

,

John A. Marwick

1

, Sonja Vermeren

1

, Adriano G. Rossi

1

, Mikala Egeblad

2

, Ian Drans

field

1

✉

&

Marc Vendrell

1

✉

Programmed cell death or apoptosis is a central biological process that is dysregulated in

many diseases, including inflammatory conditions and cancer. The detection and

quantifi-cation of apoptotic cells in vivo is hampered by the need for

fixatives or washing steps for

non-

fluorogenic reagents, and by the low levels of free calcium in diseased tissues that

restrict the use of annexins. In this manuscript, we report the rational design of a highly stable

fluorogenic peptide (termed Apo-15) that selectively stains apoptotic cells in vitro and in vivo

in a calcium-independent manner and under wash-free conditions. Furthermore, using a

combination of chemical and biophysical methods, we identify phosphatidylserine as a

molecular target of

Apo-15. We demonstrate that Apo-15 can be used for the quanti

fication

and imaging of drug-induced apoptosis in preclinical mouse models, thus creating

opportu-nities for assessing the in vivo ef

ficacy of anti-inflammatory and anti-cancer therapeutics.

https://doi.org/10.1038/s41467-020-17772-7

OPEN

1_{Centre for Inflammation Research, University of Edinburgh, EH16 4TJ Edinburgh, UK.}2_{Cold Spring Harbor Laboratory, Cold Spring Harbor, NY 11724, USA.} 3_{Institute for Molecular Bioscience, The University of Queensland, Brisbane, Queensland 4072, Australia.}4_{Instituto Biofisika (CSIC, UPV/EHU) and} Departamento de Bioquímica, Universidad del País Vasco, Campus de Leioa, 48940 Leioa, Spain.5_{Laboratory of Medicinal Chemistry and Institute of} Biomedicine U. Barcelona (IBUB), Faculty of Pharmacy, University of Barcelona, 08028 Barcelona, Spain.6_{Present address: School of Biomedical Sciences,}

Queensland University of Technology, Translational Research Institute, Brisbane, QLD 4102, Australia.7_{These authors contributed equally: Nicole D. Barth,}

Ramon Subiros-Funosas. ✉email:i.dransfield@ed.ac.uk;marc.vendrell@ed.ac.uk

123456789

P

rogrammed cell death (apoptosis) is pivotal for

main-tenance of tissues and the regulation of inflammatory

dis-eases. In contrast to necrosis, plasma membrane integrity is

preserved during apoptosis, preventing the release of intracellular

contents that can damage tissue and trigger inflammatory

responses. Tissue phagocytes recognize apoptotic cells, providing

a mechanism for the safe disposal of apoptotic material. Critically,

excessive apoptosis or failure to clear apoptotic material results in

secondary necrosis with the release of pro-inflammatory

intra-cellular contents. Therefore, the presence of apoptotic cells

represents a biomarker of the extent of tissue injury and

corre-lates to the progression, resolution, and treatment of

inflamma-tory conditions and cancer

1,2

_.

A sequence of morphological and biochemical changes occurs

during apoptosis (e.g., phospholipid exposure, caspase activation,

mitochondrial dysfunction, DNA fragmentation)

3–6

_{. Optical}

reagents for the detection of these events have been reported, but

many probes alter cellular behavior, limiting their use for

non-invasive detection of apoptosis in vivo. In particular, these

lim-itations impede studies under physiological conditions (e.g.,

intravital imaging) and in situ assessment of therapy-induced

apoptosis in preclinical models. At early stages of apoptosis, the

plasma membrane undergoes profound remodeling and the

activation of the scramblase Xkr8 promotes external exposure of

phospholipids containing phosphatidylserine (PS)-headgroups

and other phospholipids, which are normally restricted to the

intracellular leaflet

7

_{. Reagents that bind to PS exposed on the}

plasma membrane (e.g., annexins)

4,8

_{have advantages over those}

monitoring intracellular changes (e.g., caspase activation, DNA

fragmentation) because they do not require cell permeabilization

or

fixation. However, fluorescently-labeled annexins and

polarity-sensitive annexins (pSIVA) require high concentrations of free

Ca

2+

(>1 mM) to permit optimal phospholipid binding

4,5

_,

lim-iting their performance in the hypocalcemic regions that are

common in diseased tissues. Furthermore, annexins inhibit the

engulfment of apoptotic cells by phagocytes

9

_{, which precludes}

quantitative analysis of therapy-induced apoptosis in vivo.

In this study, we describe the rational design, optimization and

validation of a

fluorogenic peptide (termed Apo-15) to bind

negatively-charged phospholipids exposed on apoptotic cells.

Apo-15

behaves as a universal apoptosis probe in that it detects

apoptotic cells from multiple origins and in a broad range of

experimental conditions. Furthermore, we demonstrate that

Apo-15

enables

fluorescence imaging of apoptosis in vivo and

quan-tification of drug-induced apoptosis in two different preclinical

mouse models of acute lung injury (ALI) and breast cancer.

Results

Rational design and synthesis of

fluorogenic peptides. We

designed cyclic amphipathic peptides (termed apopeptides) which

we predicted would bind to phospholipids translocated to the outer

leaflet of apoptotic cell membranes. We focused on small cyclic

peptides because they offer several advantages: (1) resistance to

proteolytic cleavage and oxidative conditions for in vivo studies,

(2) tunability of the properties by changing the amino acid

sequence, which would allow us to optimize binding to apoptotic

cells, (3) compatibility with

fluorogenic amino acids for wash-free

imaging, and (4) smaller size than proteins for improved tissue

accessibility. Apopeptides were designed to contain combinations

of polar and hydrophobic amino acids to identify sequences

that would bind to apoptotic cell membranes and remain unable

to bind viable cells. All the apopeptides incorporated the

environmentally-sensitive Trp-BODIPY

fluorophore

10–12

_{, which}

emits bright

fluorescence after binding to provide optimal

dis-crimination between viable and apoptotic cells (Fig.

1

a, c).

A total of 15 apopeptides were prepared using solid-phase and

solution peptide synthesis (Fig.

1

b). In addition to Trp-BODIPY,

apopeptides included tryptophan (W), phenylalanine (F), leucine

(L), valine (V), and isoleucine (I) as hydrophobic residues, and

lysine (K), glutamic acid (E), and arginine (R) as polar residues. In

all cases, one glycine was included at the C-terminal end to facilitate

head-to-tail cyclization (for enhanced resistance to proteolysis

13,14

_),

and to avoid stereoisomeric mixtures. Unprotected amino acids

were used for the hydrophobic residues, whereas protected amino

acids [e.g., Fmoc-Lys(Z)-OH, Fmoc-Glu(Bzl)-OH, Fmoc-Arg

(NO

2

)-OH] were used for the polar residues. The solid-phase

peptide elongation was performed on 2-chlorotritylchloride

poly-styrene resin using conventional protocols and mild acidic cleavage

conditions [i.e., TFA:DCM (1:99)]

15

. Peptides were cyclized in a

head-to-tail fashion using COMU as the condensation reagent

16

,

and side-chain protecting groups were removed to afford all

apopeptides in high purities (>97%) after HPLC purification

(Supplementary Table 1).

The sequences and physicochemical properties of all apopeptides

are summarized in Fig.

1

b. We sought to identify those sequences

that would rapidly binding to apoptotic cells with minimal labeling

of viable cells. cLac-BODIPY, a cyclic peptide able to bind apoptotic

bodies but not apoptotic cells

11

_{, was included for comparison.}

Using a

flow cytometry-based assay, we examined the

time-dependent emission of apopeptides in mixtures of both apoptotic

and viable cells. We used human neutrophils cultured in vitro for

18 h, in which a large fraction of the cells (≥50%) undergo tissue

culture-induced apoptosis

17

_{. Apoptotic and viable populations were}

defined by positive and negative staining with AF647-Annexin V in

media containing 2 mM CaCl

2

, respectively. First, we assessed how

polar residues influenced binding by comparing two peptides with

similar molecular weight and clog P (Supplementary Table 1 and

Fig.

1

b), but with either negatively-charged (Apo-0) or

positively-charged (Apo-2) residues. We chose glutamic acid (E) as a

negatively-charged amino acid over aspartic acid to avoid synthetic

complications due to the potential formation of aspartimides

18

_.

Apo-2

showed selective binding to apoptotic cells over viable cells

when compared with Apo-0, indicating the importance of positive

charges for binding to negatively-charged phospholipids on

apoptotic cell membranes. Next, we generated amphipathic peptides

containing positively-charged amino acids and other residues that

would alter binding to apoptotic cell membranes

19,20

_{. Specifically,}

we synthesized apopeptides to examine the influence of (1)

aromatic vs non-aromatic hydrophobic residues (Apo-3, 4, and

Apo 9

–10), (2) alternate vs sequential charges (Apo 5–8), and (3)

overall polarity as determined by clog P values (Apo 11–14).

Temporal analysis indicated that recognition of apoptotic cells

occurred rapidly, with most apopeptides showing

≥80% of full

binding in <4 min (Supplementary Table 2). From the screening,

we quantified parameters that defined the selectivity and affinity of

apopeptides: (1) preferential binding to apoptotic vs viable cells as

fluorescence fold increase (Ff), (2) background fluorescence on

viable cells (Bf), and (3) retention of binding upon washing (Rt)

(Fig.

1

b). Several apopeptides showed good discrimination between

apoptotic and viable cells (Ff

≥ 10: Apo-2, 3, 6, and 8), negligible

fluorescence on viable cells (Bf ≤ 3: Apo-0, 4, 11, 12, 13, and 14)

and reasonable resistance to washing (Rt

≥ 40%: Apo-2, 3, 6, 8, and

11). Overall, peptide polarity or clog P was related to retention of

labeling (Fig.

1

b). Although markedly polar peptides (clog P <

−4,

Apo-13

and Apo-14) bound with fast kinetics, their signal was lost

after washing. In contrast, less polar peptides (clog P >

−1, Apo-11

and Apo-12) displayed slower binding rates but their binding was

relatively resistant to washing. These analyses suggested that

apopeptides with balanced polarity (clog P between

−1 and −4)

exhibited better labeling. Apo-8 presented the highest retention of

signal but also showed the highest binding to viable cells. Our

NH NH NH H N NH HN H N R1 R3 R5 R2 R4 O O O O_O O N H O N B N F F i) Cleavage (TFA:DCM 1:99) Cl N H N B N H N N H O O O O N H O R5 H N O R4 N H R3 O H N R2 H2N R1 O ii) Cyclization (COMU, DIPEA) F F Fmoc SPPS NH NH NH H N NH HN H N R1 R3 R5 R2 R4 O O O O O O N H O N B N F F CTC-PS Resin Oxyma Pure DIC iii) Deprotection NON FLUORESCENT Apoptotic HIGHLY FLUORESCENT NH NH NH H N NH HN H N R1 R3 R5 R2 R4 O O O O O O N H O N B N F F Turn-on Fluorescence (520 nm) Viable

a

b

Fluorescence (R FU) Apo-0 Bf: x y x y Apoptotic Viable 0 104 103 102 101 0 104 103 102 101 60 120 180 0 Apoptotic Viable Ff: x' y' y' x' Time (s) 60 120 180 0 Apo-15

c

d

Nuclear Counterstain AF647-Ann V Counterstain Nuclear Counterstain AF647-Ann V Counterstain Apo-0 Apo-15

e

24 h PBS 24 h protease 24 h H2O2 HPLC purity (Apo-15) 98 ± 2% 95 ± 2% 96 ± 1% Environmentally Sensitive Reporter Apopeptide Binding Sequence

Peptides R1–R5 clog P Fold increase (Ff)

[apoptotic vs viable] Background (Bf) [viable cells] Retention (Rt, % ) Apo-0 EEEWW –2.97 –2.78 –2.93 –3.08 –2.78 –2.78 –2.78 –2.78 –3.41 –3.94 –0.68 1.43 –4.63 –5.38 –3.38 –5.49 <1 2.4 6.6 4.7 3.0 3.6 5.6 4.0 11.4 n.d. n.d. 1.2 1.3 0.8 0.9 2.4 4.6 n.a. Apo-2 KKKWW 18 62 Apo-3 KKKWF 14 47 Apo-4 KKKFF 6 15 Apo-5 KWKWK 7 8 Apo-6 KKWWK 18 40 Apo-7 WWKKK 8 22 Apo-8 KWWKK 13 99

Apo-9 KKKLI <1 n.a.

Apo-10 KKKVI <1 n.a.

Apo-11 KKWWW 7 47

Apo-12 KWWWW <1 n.a.

Apo-13 KKKKW 6 9

Apo-14 KKKKK <1 n.a.

Apo-15 RKKWF 14 42

cLac-BODIPY Reference 11 <1 n.a.

Apo-0 to 15

Fig. 1 Design, synthesis, and in vitro screening of apopeptides. a Synthetic scheme for the preparation of apopeptides (CTC-PS: 2-chlorotritylchloride polystyrene resin). Stability analysis ofApo-15 (50μM) under different proteolytic and oxidative environments. b Sequences, physicochemical and binding properties of apopeptides and cLac-BODIPY. In vitro screening of the peptides (100 nM) was performed by real-timeflow cytometry in mixtures of apoptotic (AF647-Annexin V positive) and viable (AF647-Annexin V negative) neutrophils (_{n ≥ 3, biologically independent experiments). c Fluorescence} activation of apopeptides in apoptotic (right) over viable cells (left), with Trp-BODIPY as the environmentally-sensitive reporter. Pictograms ofApo-15 under 365 nm light excitation.d Representativeflow cytometry plots of Apo-0 and Apo-15 binding to mixtures of apoptotic (highlighted in green dots) and viable cells (highlighted in black dots). Fluorescence emission was recorded for around 4 min after addition of apopeptides (100 nM) without washing. Backgroundfluorescence (Bf) was calculated as the ratio of fluorescence recorded before (x) and 2 min after (y) peptide addition. Fold increase (Ff) in apoptotic cells was calculated as a ratio offluorescence recorded for apoptotic cells (y′) to that of viable cells (x′) after 2 min. e Representative fluorescence confocal microscopy images (from three independent experiments) of viable (white arrows) and apoptotic cells (yellow arrows) after incubation withApo-0 (left, green) and Apo-15 (right, green) (both at 100 nM). Cells were co-stained with Hoechst 33342 (7_{μM, blue) and} AF647-Annexin V (5 nM, red) as nuclei and apoptosis markers, respectively (λexc.: 405, 488, 633 nm:λem.: 450, 525, 670 nm). Scale bars: 10μm. Source data

analyses also revealed the importance of non-electrostatic

interac-tions, with apopeptides lacking hydrophobic aromatic residues

(Apo-9, 10, and 14) exhibiting poor retention of labeling. Besides,

among aromatic amino acids, tryptophan increased specificity

when compared with phenylalanine (Apo-2 vs Apo-4).

Considering all these results, we decided to further optimize the

Apo-3

sequence (Ff: 14; Bf: 4.7; Rt: 47%) to reduce its background

fluorescence on viable cells while retaining full binding to

apoptotic cells. Arginine-rich peptides have been described to

strongly bind molecular targets in other amphipathic sequences,

partially due to the larger polar surface area of arginine compared

with lysine

21

_{. Therefore, we synthesized Apo-15 by replacing one}

lysine with an arginine residue. Apo-15 [sequence: c(RKKWFW

(BODIPY)G)] displays high selectivity for apoptotic cells (Ff: 14),

marginal background

fluorescence on viable cells (Bf: 2.4) and

good retention of signal after three washes with PBS (Rt: 42%).

Representative

flow cytometry plots for Apo-0, which does not

bind to apoptotic cells, and Apo-15 are shown in Fig.

1

d. We also

examined Apo-0 and Apo-15 for imaging apoptotic cells in the

presence of viable cells (Fig.

1

e). Apo-15 displays

fluorescence in

the green region of the visible spectrum, being compatible with

conventional GFP and FITC

filters (λ

abs.

: 500 nm;

λ

em.

: 530 nm;

Supplementary Fig. 1). Notably, Apo-15 displays around 10-fold

brightness (25,000 M

−1

cm

−1

) over other

environmentally-sensitive probes for apoptosis [e.g., pSIVA (2500 M

−1

cm

−1

), N,

N′-didansyl-L-cystin (3000 M

−1

_cm

−1

_{) (Table}

₁

_{)], which allows}

the staining of apoptotic cells at nanomolar concentrations

(30–100 nM, Supplementary Fig. 2). Apo-15 also exhibits high

chemical stability under proteolytic and oxidative conditions.

HPLC analysis confirmed >95% of Apo-15 remained stable after

incubation with a protease cocktail or the oxidizing agent H

2

O

2

(Fig.

1

a and Supplementary Figs. 3, 4). Altogether, these properties

made Apo-15 an optimal candidate for further characterization.

Apo-15 delineates apoptotic cells in diverse environments.

Next, we evaluated Apo-15 for the general detection of apoptotic

cells from different species and lineages. We observed that

Apo-15

selectively stained apoptotic cells regardless of their origin.

Specifically, we examined myeloid cells (neutrophils, both human

and mouse, Supplementary Fig. 5), lymphoid cells (BL-2, Burkitt

lymphoma) and primary epithelial cells. We performed these

experiments in the presence of AF647-Annexin V to corroborate

that Apo-15 stains apoptotic and not viable cells. Notably, we

observed very similar staining for Apo-15 and AF647-Annexin V

in media containing 2 mM CaCl

2

(Fig.

2

a, b). Furthermore,

Apo-15

labeling proved to be independent of the method used to

induce apoptosis [e.g., myeloid: tissue culture-induced apoptosis

by culture at 37 °C for 18 h; lymphoid: irradiation with a CL-1000

Ultraviolet Crosslinker UVP at 254 nm; epithelial: treatment with

staurosporine (1 µM) for 6 h], which highlights the compatibility

of Apo-15 with multiple experimental conditions.

A limitation of annexins is their dependence on high

concentrations of free Ca

2+

(>1 mM), which affects their use in

hypocalcemic environments in diseased tissues

22

_{. Therefore, we}

decided to assess whether Apo-15 was able to delineate apoptotic

cells independently of the concentration of free divalent cations.

Notably, we observed robust binding of Apo-15 to myeloid and

lymphoid apoptotic cells in the presence of the divalent cation

chelator EDTA (2.5 mM), whereas AF647-Annexin V failed to

bind under the same experimental conditions (Fig.

2

b–d). Ca

2+

-dependent binding to apoptotic cells was also observed for

polarity-sensitive annexins (pSIVA)

8

(Supplementary Fig. 6). The

divalent cation-independence of Apo-15 represents a major

advantage over annexins and allows direct monitoring of

apoptosis in most conditions likely to be encountered in vivo.

Table

1

Comparative

analysis

of

Apo-15

and

commercially

-a

vailable

fl

uorescent

probes

for

the

detection

of

apoptosis.

Probe Apo-15 Annexins pSIVA N,N ′-Didansyl-L-cystin JC1 Nonyl acridine orange Active caspase-3/7 PI Fluorescence wavelengths (λexc. /λ em. , nm) 500/520 Variable 465/540 365/530 488/ 530 –590 495/519 Variable 535/617 Brightness (φ × ε) 25,000 Variable 2500 3000 n.a. 20,000 Variable <100 Molecular weight 1.3 kDa ~36 kDa ~35 kDa 707 Da n.a. 473 Da Variable 668 Da Molecular target PS PS PS n.a. Mito memb potential Cardiolipin Activated caspases 3/7 DNA Early /late apoptosis Early Early Early Early Mid Mid-late Late Late Divalent cation-independence Yes No No Yes Yes Yes Yes Yes Wash-free imaging Yes High Ca 2+ High Ca 2+ Yes No No No No Apoptotic cell sorting Yes High Ca 2+ High Ca 2+ No No No No No Functionally neutral Yes No No n.a. No n.a. No Yes Phagocytic engulfment marker Yes No No No No No No No Applications FC, IHC, CM, FP, intravital FC, IHC, CM (high Ca 2+ ) FC, IHC, CM (high Ca 2+ ) FC, CM FC, CM FC, CM FC, IHC FC, IHC References This work 4 , 59 8 , 60 19 , 61 5 , 62 63 , 64 3 , 65 66 , 67 CM confocal microscopy ,FC fl ow cytometry, FP fl uorescence polarization, IHC immunoh istochemistry, PI propidium iodid e.

To confirm that cells stained by Apo-15 under divalent

cation-free conditions were apoptotic, we sorted a mixed population of

apoptotic and viable human neutrophils (gating strategy in

Supplementary Fig. 7) according to Apo-15 labeling and examined

their morphology by microscopy. Apo-15-positive neutrophils

exhibited a pyknotic nucleus characteristic of chromatin

con-densation and degradation, together with cell shrinkage, which are

hallmarks of apoptosis (Fig.

2

c)

23

_{. In contrast, Apo-15-negative}

cells showed the typical multilobed nuclear morphology of viable

neutrophils (Fig.

2

c). The morphological appearance of

Apo-15-positive cells was examined by scanning electron microscopy.

Apo-15-positive neutrophils showed evidence of

apoptosis-associated blebbing and pitting of the plasma membrane

(Supplementary Fig. 8)

24,25

_{. On the other hand, the presence of}

microvilli-like structures, which are typical of viable cells, were

observed on the plasma membrane of Apo-15-negative cells

(Supplementary Fig. 8). Apo-15 also discriminated between

different types of cell death by

fluorescence lifetime imaging

(FLIM). Human BL-2 cells were induced into apoptosis or

necrosis by differential UV irradiation, incubated with Apo-15

and imaged under a FLIM microscope to reveal that apoptotic and

necrotic cells could be discriminated by their

fluorescence lifetimes

(Supplementary Fig. 9). Altogether, these results demonstrate that

Apo-15

is a generic marker of apoptotic cells under different

physiological environments and compatible with multiple

biolo-gical studies.

Apo-15 rapidly binds PS for real-time and wash-free imaging.

In view of the specific labeling of apoptotic cells by Apo-15, we

examined whether the amphipathic nature of Apo-15 conferred

Apo-15 Merged (+Hoechst)

Myeloid L ymphoi d Epithelial AF647-Annexin V AF647-Annexin V Apo-15 AF647-Annexin V FACS sorting Percentage stained cells (% ) p < 0.0001 p < 0.0001 p < 0.0001 p = 0.6738 p = 0.0505 p = 0.0776 Apo-15

a

b

d

c

AF647-Annexin V Apo-15 103 102 101 101 ₁₀2 ₁₀3 AF647-Annexin V ₀ 103 102 101 101 102 103 0 Apo-15 p = 0.0125 p = 0.7129 t (h) UV (mJ cm–2 ) Ca2+ (2 mM) Apo-15 Annexin V

0 0 18 18 18 18 n.a. n.a. n.a. n.a.

n.a. n.a. n.a. n.a. n.a. n.a. 0 0 300 300

+ + + + – + + + + + – + – + – + – + – + – – + – + – + – + – n.a. n.a. 300 300 + – – + – – Staurosp. Ca2+_{(2 mM)} Apo-15 Annexin V + + + + – + – + – + – + – – + – + – + + + + – – Percentage stained cells (% ) p < 0.0001 p = 0.7563 p = 0.0016 p = 0.040 +2 mM Ca2+ (annexin binding) 103 102 101 101 ₁₀2 ₁₀3 0 103 102 101 101 ₁₀2 ₁₀3 0 +2 mM Ca2+ (annexin binding) +2.5 mM EDTA (annexin not binding) +2.5 mM EDTA (annexin not binding)

Lymphoid Myeloid Epithelial 0 10 20 30 0 20 40 60 80 100

Fig. 2 Apo-15 binds to apoptotic cells of different origin in multiple environments. a Representativefluorescence confocal microscopy images (from three independent experiments) human apoptotic (yellow arrows) and viable (white arrows) cells from different lineages: BL-2 (lymphoid), neutrophils (myeloid), and primary airway epithelial cells (epithelial). Cells were incubated withApo-15 (100 nM, green), AF647-Annexin V (5 nM, red), and Hoechst 33342 (7_{μM, blue) for 10 min and imaged under a fluorescence confocal microscope (λ}exc.: 405, 488, 633 nm:λem.: 450, 525, 670 nm). Scale bars: 10µm.

b Divalent cation-independent binding of Apo-15 to apoptotic cells of different origin (top plots: lymphoid cells, bottom plots: myeloid cells). Mixtures of apoptotic (highlighted in green dots) and viable cells (highlighted in black dots) were stained with AF647-Annexin V (25 nM) andApo-15 (100 nM) in the presence of 2 mM CaCl2(left) or 2.5 mM EDTA (right). Representative histograms showingApo-15 binding (x-axis) vs AF647-Annexin V binding (y-axis)

acquired on 5L LSRflow cytometer (n = 5). c Cells were sorted on their Apo-15 positivity (green populations in panel b) or negativity (black populations in panelb). Gating strategy in Supplementary Fig. 7. Morphological analysis (four independent images from two independent experiments) of cytocentrifuge preparations ofApo-15-positive and Apo-15-negative cells under bright_{field microscopy (scale bar: 10 µm). d Quantification of fluorescence staining of} neutrophils (myeloid,n ≥ 5), BL-2 cells (lymphoid, n ≥ 3) and primary airway epithelial cells (epithelial, n = 5) before and after induction of apoptosis and upon treatment withApo-15 and AF647-Annexin V. Data acquired on 5 L LSRflow cytometer and presented as mean values ± SEM. P values obtained from two-tailedt tests. Source data (in d) are provided as a Source data file.

binding to negatively-charged phospholipids (e.g., containing

PS-headgroups) that are exposed on the plasma membrane of

apoptotic cells but are inaccessible in most viable cells. We used

giant unilamellar vesicles (GUVs, 1–10 μm) composed of neutral

phospholipids (containing only phosphatidylcholine

(PC)-head-groups, 0.2 mM) or mixed with negatively-charged

PS-phospho-lipids

(PC:

0.14 mM,

PS:

0.06 mM).

Using

fluorescence

microscopy, we observed that Apo-15 stained PC:PS GUVs with

brighter intensity than PC GUVs of similar size (Fig.

3

a, b).

Binding to PS in PC:PS GUVs was confirmed by co-staining with

the positive control AF647-Annexin V in 2 mM CaCl

2

. On the

other hand, we did not observe significant differences in

fluores-cence intensity between PC:PS GUVs and PC GUVs when stained

with the lipid marker Lissamine-Rhodamine DOPE. Furthermore,

we analyzed the specificity of Apo-15 for PS over other

phos-pholipids [i.e., cardiolipin, PC, phosphatidylglycine,

phosphati-dylethanolamine (PE)] and also phosphatidic acid. Apo-15

exhibited dose-dependent and significantly stronger binding to PS

(Fig.

3

b and Supplementary Fig. 10) and showed preferential

binding to PS over PE, which is also known to localize on the

outer leaflet of the plasma membrane following induction of

apoptosis

26

. Apo-15 proved compatible with anisotropy readings,

104 103

Time (apoptosis progression)

Apo-15 AF647-Ann V Time (min) 0' 4' 13 ' 0' 9' Apo-15 Fluorescence Lissamine Apo-15 AF647-Ann V Merged

PC PC:PS

b

a

d

f

e

g

c

p-values PS vs. PG: 0.0040 PS vs. PE: 0.0017 PS vs. CL: 0.0019 PS vs. PA: 0.0048 PS vs. PC <0.0001 BL-2 (PS exposure) PLB-985 (lack of PS exposure) 40 ± 3 % Apo-15+ 1.0 ± 0.3 % Apo-15+ 0 3.6 ± 1 % Apo-15+ 4 ± 1 % Apo-15+ 104 104 103 103 102 102 101 101 0 104 103 102 101 AF647-A nnexin V AF647-A nnexin V Apo-15 Fluorescence 0 Viable Apoptotic Counts [Annexin V] no Annexin 5 nM 20 nM 85 nM_{270 nM} 5 nM 20 nM 85 nM 270 nM p = 0.0006 p = 0.0458 Apo-15 only PS PG PE CL PA PC 4000 8000 12,000 16,000 Fluorescence Apo-15 (R FU) 0 2000 4000 6000 8000

Fluorescence Apo-15 (RFU

) 0 0 200 400 Time (s) 600 800 1000 10,000 20,000 30,000 40,000 Fluorescence intensity (RFU )

showing concentration-dependent

fluorescence polarization in

PS-containing liposomes due to the environmentally-sensitive nature

of Trp-BODIPY (Supplementary Fig. 11).

Next, we examined whether Apo-15 could bind PS in cell

membranes. First, we performed competitive assays between

AF647-Annexin V and Apo-15 at different concentrations in

co-cultures of viable and apoptotic cells. We observed that high

concentrations of AF647-Annexin V could reduce binding of

Apo-15

to apoptotic cells (Fig.

3

c) and also that Apo-15 reduced

Annexin V binding (Supplementary Fig. 12), suggesting that

Annexin V and Apo-15 compete for binding sites on the surface

of apoptotic cells. Secondly, we used a human cell line (i.e.,

PLB-985) that is unable to translocate phospholipids to the outer

leaflet of the plasma membrane during apoptosis due to lack of

scramblase Xkr8

27

_{. As a control, we used human BL-2 cells,}

which expose PS on their membrane upon induction of apoptosis.

Apoptosis in both PLB-985 and BL-2 cells was induced by

treatment with staurosporine, followed by incubation with

Apo-15

and

flow cytometry analysis. Apo-15 stained around 40% of

BL-2 cells, which were also stained with AF647-Annexin V,

confirming that they were apoptotic (Fig.

3

d). In contrast, only

1% of PLB-985 cells were Apo-15-positive under the same

experimental conditions (Fig.

3

d). We corroborated that the

treatment with staurosporine induced apoptosis in PLB-985 cells

by detection of cleaved caspase-3/7 (Supplementary Fig. 13)

28,29

_.

Taken together, these results indicate that the selective binding of

Apo-15

occurs at early stages of apoptosis when PS is

translocated to the outer leaflet of apoptotic cells.

Given the rapid binding and

fluorogenic properties of Apo-15,

we evaluated its application for real-time imaging of cells actively

undergoing apoptosis. BL-2 cells were induced to apoptosis by

exposure to UV light and treated with Apo-15 just before

time-lapse movies were acquired under wash-free conditions in a

live-cell spinning-disk microscope. We observed minimal background

fluorescence suggesting that the Apo-15 is compatible with

wash-free imaging with similar signal-to-noise ratios to annexin-based

reagents (Supplementary Fig. 14). We also observed cells that

transitioned from being non-fluorescent (i.e., viable) to

Apo-15-labeled (i.e., apoptotic), confirmed by simultaneous staining with

AF647-Annexin V (Fig.

3

e and Supplementary Movies 1 and 2).

Furthermore, time-lapse quantification of the fluorescence

intensity in multiple transitioning cells demonstrated that

Apo-15

bound rapidly to apoptotic cells, with more than 90% of the

association occurring in <10 min (Fig.

3

f). Next, we exploited

these features of Apo-15 to non-invasively image the dynamics of

formation of apoptotic bodies and vesicles, which have a critical

role in intercellular communication

30

_{, and are difficult to image}

with annexins due to the formation of two-dimensional crystal

lattices that alter membrane dynamics

31

_{. Time-lapse imaging}

following induction of apoptosis in the presence of Apo-15

revealed rapid release (<10 min) of subcellular vesicles and, in

some cases, reintegration of vesicles back into the plasma

membrane (Fig.

3

g and Supplementary Movie 3). These results

confirm the utility of Apo-15 for wash-free imaging of early

apoptotic events at both cellular and subcellular levels.

Apo-15 enables imaging of apoptotic cell clearance. Functional

neutrality is an essential requirement for probes that enable

imaging of biological processes under live-cell conditions. First,

we tested whether Apo-15 affected the rate at which apoptosis

occurs in vitro. For these experiments, we cultured human

neu-trophils in the presence of different concentrations of Apo-15 and

examined the extent of tissue culture-induced apoptosis at

dif-ferent timepoints. Even when used at higher concentrations than

those required for imaging or

flow cytometry, Apo-15 did not

affect the detection or the quantification of apoptosis, as

mea-sured by staining with AF647-Annexin V (Fig.

4

a).

One potential limitation to the use of annexins for imaging

apoptosis is their interference with the clearance of apoptotic cells

by phagocytes. This is important for in vivo imaging studies, since

the perturbation of apoptotic cell clearance could result in over- or

under-estimation of the extent of apoptosis within a tissue. We

examined whether Apo-15 affected the engulfment of apoptotic

cells using a well-established in vitro assay to quantify the extent of

phagocytosis in human monocyte-derived macrophages (MDMs)

in the presence of the PS-opsonin protein S (Fig.

4

b)

32,33

_{. Unlike}

Annexin V, which significantly reduces (~50%) the phagocytic

capacity of MDMs, Apo-15 did not inhibit the phagocytosis of

apoptotic cells, even when it was used at high concentrations

(Fig.

4

c). These observations were further corroborated by

fluorescence microscopy assays in co-cultures of human MDMs

and apoptotic neutrophils, where we clearly observed engulfment

of Apo-15-labeled apoptotic cells and retention of Apo-15 staining

after phagocytosis (Fig.

4

d). This result opens the possibility of

using Apo-15 as a probe for tracking phagocytosis under live-cell

conditions.

Apo-15 detects drug-induced apoptosis in mouse models.

Preclinical and clinical studies have shown that low levels of free

Ca

2+

are common in disease

22

, with many studies (e.g., intravital

imaging, in vivo therapy evaluation) being hampered by the lack

of specific apoptosis reagents that are effective in those

physio-logical environments. Given the rapid, specific and divalent

Fig. 3 Apo-15 rapidly binds exposed PS on the surface of apoptotic cells. a Fluorescence microscope images (three independent images from two independent experiments) of GUVs with differential lipid content (PC and PC:PS). Vesicles were stained with 0.3 mol% Lissamine-Rho-DOPE (blue), Apo-15 (100 nM, green) and AF647-Annexin V (5 nM, red) (λexc.: 488, 561, 633 nm:λem.: 525, 593, >650 nm). Scale bar: 10µm. b Binding specificity of

Apo-15 upon incubation with different phospholipids (PS: phosphatidylserine, PG: phosphatidylglycine, PE: phosphatidylethanolamine, CL: cardiolipin, PA: phosphatidic acid, PC: phosphatidylcholine). Data acquired on a spectrophotometer (λexc.: 450 nm,λem.: 520 nm,n > 10, except for PA) and presented as

mean values ± SEM.c Representative histograms showing Apo-15 (100 nM) labeling of mixed populations of viable and apoptotic neutrophils upon competition with increasing concentrations of AF647-Annexin V, together with quantification of the mean fluorescence intensity of Apo-15-stained apoptotic cells (n ≥ 3) presented as mean values ± SEM. d Representative histograms of BL-2 and PLB-985 cells after treatment with staurosporine (1 µM for 3 h) and staining with AF647-Annexin V andApo-15 (n = 3). Apo-15-stained cells are highlighted as green dots and Apo-15-negative cells are highlighted as black dots. Gating strategy in Supplementary Fig. 7.e Time-lapsefluorescence snapshots (from three independent experiments) of apoptotic BL-2 cells after UV irradiation (300 mJ cm−2) (Supplementary Movies 1 and 2). BL-2 cells were treated withApo-15 (green) and AF647-Annexin V (red) and imaged every 30 s for 15 min without washing. Arrows point at a cell when viable (white) and when undergoing apoptosis (yellow). Scale bar: 10µm. f Quantification of Apo-15 fluorescence intensity and presented as mean values ± SEM over time from Supplementary Movie 1 (n = 3 independent measurements).g Time-lapse_{fluorescence images (from five independent cells) of subcellular apoptotic bodies (white arrow) in apoptotic neutrophils} stained withApo-15 (green). Images were acquired every 1 s for 15 min (Supplementary Movie 3). Scale bar: 2µm. For (b) and (c), P values were obtained from two-tailedt tests. Source data (in b, c, and f) are provided as a Source data file.

cation-independent binding of Apo-15, together with its

wash-free capabilities and functional neutrality, we examined whether

Apo-15

could be used for imaging of apoptotic cells in vivo in a

mouse model of ALI. This model allows temporal control

of neutrophil recruitment by intratracheal administration of

lipopolysaccharide (LPS), which is followed by neutrophil

apop-tosis and subsequent clearance by phagocytic macrophages

34

_.

Importantly, this is a preclinical model in which the extent of

apoptosis can be modulated by cyclin-dependent kinase

inhibi-tors (CDKi)

35

_{. CDKi have profound effects on inflammatory cells}

and some are under development as anticancer therapies

36,37

.

First, we confirmed that Apo-15 could detect CDKi-induced

apoptosis in vitro. Neutrophils incubated with the CDKi

R-roscovitine exhibited increased apoptosis by

flow cytometry (as

defined by Annexin V-positive staining) with a concomitant

increase in the proportion of Apo-15 stained cells (Fig.

5

a).

Apo-15

staining of CDKi-induced apoptosis was also

caspase-dependent as confirmed by a significant reduction in the

fluorescent signal after treatment with the caspase inhibitor

QVD

34

_(Fig.

₅

_{a). Next, we induced ALI in vivo in mice by}

intratracheal administration of E. coli-derived LPS (1

μg per

mouse), and the apoptotic load in the lungs relative to saline

control was increased by intraperitoneal administration of CDKi

AT7519

38

_(Fig.

₅

_{b). In all mice, we administered Apo-15 in vivo}

and, 30 min later, we collected bronchoalveolar lavage

fluid

(BALF) together with lung tissues. BALF and lung slices from

LPS-only and LPS

+ CDKi mice were analyzed by

multipara-meter

flow cytometry to determine the proportion of apoptotic

cells that had been stained by Apo-15 in vivo (gating strategy in

Supplementary Fig. 15). Notably, a proportion of CD45

+ Ly6G+

a

d

c

0 20 40 60 80

Percentage apoptotic cells (%

) 3 h 18 h p = 0.9724 p = 0.0040 p = 0.6814 Protein S Apo-15 Ann V – – – p = 0.0008 p = 0.0011 p < 0.0001 – – + – + – – + + 20 40 60 80 Percentage phagocytosis (% ) 60 nM 600 nM

b

phago MDMs non-phago MDMs 0 105 104 103 50 150 250 50 150 250 FSC (×103) Fluorescence pHrodo no Apo-15 60 nM Apo-15 600 nM Apo-15 62% 67% 63% +

Fig. 4 Apo-15 does not interfere with the phagocytosis of apoptotic cells. a Percentages of human neutrophils undergoing apoptosis, as indicated by AF647-Annexin V staining, induced after culture at 37 °C for different times (3 h: left; 18 h: right). Neutrophils were cultured in the absence (black) or presence ofApo-15 (gray: 100 nM, white: 300 nM). Data presented as mean values ± SEM (_{n = 3). b Representative histograms of glucocorticoid-treated} human MDMs after phagocytosis of pHrodoTM_{-labeled neutrophils in the presence of different concentrations ofApo-15. Cells were analyzed using a 5L} LSR cytometer with MDM identified based on their distinctive forward and side scatter properties (n = 4). Gating strategy in Supplementary Fig. 15A. c Flow cytometric quanti_{fication of phagocytosis of pHrodo}TM_{-labeled neutrophils by human MDMs. Human MDMs and neutrophils were co-cultured in} the absence or presence of protein S (25 nM) andApo-15 (60 or 600 nM) or AF647-Annexin V (25 nM). Data presented as mean values ± SEM (n ≥ 3). d Representative brightfield and fluorescence microscope images (from four independent experiments) of Apo-15-treated human neutrophils (green) in co-culture with human MDMs. Red arrows identifynegative viable neutrophils, and yellow arrows indicate MDM that have engulfed Apo-15-labeled apoptotic neutrophils. Scale bar: 20µm. For (a) and (c), P values were obtained from two-tailed t tests. Source data (in a and c) are provided as a Source datafile.

neutrophils in BALF were stained with Apo-15 (Fig.

5

d).

Furthermore, we quantified the effects of CDKi-induced

apop-tosis in vivo by measuring the percentages of apoptotic

neutrophils in total cell counts. BALFs from LPS

+ CDKi-treated

mice showed a significantly larger population of Apo-15-labeled

cells than BALFs from LPS-treated mice (29% vs. 5%, Fig.

5

d).

We confirmed that CD45 + Ly6G+ cells labeled by Apo-15

in vivo were apoptotic by co-staining with AF647-Annexin V

ex vivo (Fig.

5

d). Next, we acquired

fluorescence microscopy

images of lung tissue sections from mice under different regimes

(Fig.

5

c, e). We observed increased numbers of Apo-15-positive

cells in lungs from LPS

+ CDKi-treated mice when compared

with lungs of LPS-treated mice (90 vs 32 cells per

μm

2

_tissue,

Fig.

5

c) and with lungs from mice that had not been treated with

Apo-15

(Supplementary Figs. 16 and 17). Notably, the signal of

Apo-15-labeled cells was stable following processing with 10%

formalin, suggesting compatibility with

fixed tissue samples.

Furthermore,

flow cytometric analysis demonstrated that Apo-15

could stain CD45

+ CD11c + Annexin V− phagocytic

macro-phages in vivo (gating in Supplementary Fig. 15 and results in

Supplementary Fig. 18). Since viable macrophages were not

stained when BALFs were incubated with Apo-15 ex vivo

(Supplementary Fig. 18), we suggest that these macrophages

acquire

fluorescence because of internalization of Apo-15-labeled

cells, further highlighting the potential utility of Apo-15 for the

identification of cell populations that have phagocytosed

apoptotic cells.

We used Apo-15 for intravital imaging of mice undergoing

ALI using optical windows providing direct access to lungs in

living mice

39

. Following direct intratracheal administration of

Apo-15, we performed

fluorescence imaging in vivo to compare

the signals in the lungs of healthy and ALI mice. In healthy,

saline-injected control mice, we observed negligible Apo-15

staining whereas we observed bright stable staining in the lungs of

mice undergoing ALI (Fig.

5

f). Furthermore, we examined the

staining of Apo-15 following drug-induced apoptosis in the

Apo-15 + neuts (% ) LPS+ CDKi 10 30 40 0 20 LPS LPS+CDKi LPS only LPS +CDKi Apo-15 + cells per 0.25 µµ m 2 section 40 120 160 80 LPS only LPS+CDKi

d

e

c

f

g

p = 0.0068 p < 0.0001 Saline

a

Apo-15-labeled cells (% ) CDKi QVD – – + + + 20 40 60 80 +

b

p = 0.0015 p = 0.0025

ALI model Tumour model

DAPI Apo-15 Casp-3

t (h) 0 18 36 37–48 LPS (1 g)µ CDKi Apo-15 (5 M)µ Harvest, imaging Cell recruitment Resolution of Inflammation 101 ₁₀2 ₁₀3₁₀4 ₁₀1 ₁₀2 ₁₀3 ₁₀4 104 103 102 101 0 Apo-15 fluorescence Ex vivo Annexin V LPS LPS LPS+CDKi LPS (1 µg) DAPI+Apo-15 DAPI+Apo-15

Fig. 5 In vivo administration of Apo-15 labels drug-induced apoptosis in different mouse models. a Human neutrophils were treated withR-roscovitine (20µM) in the absence or presence of QVD (10 μM). Apo-15-positive cells were determined by flow cytometry (n = 4). b Experimental protocol for acute lung injury: LPS (20_{μg mL}−1) was instilled intratracheally and mice received CDKi (30 mg kg−1) or saline intraperitoneally 18 h later.Apo-15 was administered intratracheally at 36 h.c Double-blind quantification of Apo-15-positive cells in lung tissue from mice that received LPS or LPS and CDKi (n = 8 per group).d Flow cytometric analysis of BALF from mice that received LPS (20μg mL−1) or LPS (20μg mL−1) and CDKi (AT7519, 4 mg mL−1). Both groups were administeredApo-15 (5_{μM) in vivo and incubated with AF647-Annexin V (25 nM) ex vivo. Data presented as mean percentages ± s.d. of} Apo-15-positive CD45+ LY6G+ apoptotic neutrophils (n = 4). Representative dot plots of CD45 + LY6G+ neutrophils with Apo-15 positive cells highlighted in green andApo-15-negative cells in black. Gating strategy in Supplementary Fig. 15B. e Fluorescence microscope images of sections of lung tissues from mice that received LPS or LPS and CDKi. Yellow arrows identifyApo-15-positive events (green) from images withn = 8 per group. Scale bar: 100µm. f Low-magnification intravital images of lungs of saline-treated and LPS-treated mice (LPS: 20 μg mL−1).Apo-15 was administered in vivo (5μM, green) and nuclei were counterstained with DAPI (18μM, blue) (λexc.: 405, 488 nm;λem.: 450, 525 nm). Representative pictures fromn = 3 per group.

Scale bar: 50μm. g High-magnification fluorescence images (from three independent experiments) of tumors from xenograft breast cancer mouse models after treatment with cisplatin (10_{μg g}−1).Apo-15 was administered i.v. (5_{μM, green) and nuclei were counterstained with DAPI (10 μM, blue) (λ}exc.: 405,

488 nm;λem.: 450, 525 nm). Apoptotic cells were stained with anti-active caspase-3 (300 ng mL−1, yellow). Scale bar: 50μm. For (a), (c), and (d), p values

widely used MMTV-PyMT breast cancer mouse model, which

spontaneously develops tumors in the mammary glands. After

tumors were grown to a size between 0.5 and 1 cm, mice were

treated with cisplatin (10

μg g

−1

) to induce apoptosis and Apo-15

was injected in vivo intravenously via tail vein. As shown in

Fig.

5

g, the signal of Apo-15 was retained in tumor tissues after

OCT embedding and sectioning, which allowed us to compare its

localization with that of active caspase-3. These results confirmed

that Apo-15 can label apoptotic cells in vivo as defined by activity

of caspase-3, and that it can be administered in vivo via different

routes and under various experimental conditions.

Discussion

Dysregulation of apoptosis is a defining feature of the progression

of inflammatory diseases and of cancer. Although a number of

probes for the detection of apoptosis using non-optical imaging

modalities (e.g., positron emission tomography, single-photon

emission computed tomography) have been described

40,41

_{, there}

are very few optical probes that can be used in vivo (Table

1

). In

particular, chemical degradation in an inflammatory

micro-environment (e.g., protease or oxidative), dependence on in vivo

availability of high concentrations of free divalent cations and

perturbation of cellular functions represent key limitations.

We have rationally designed and synthesized

fluorogenic

heptapeptides that specifically overcome these limitations,

allowing us to identify a highly stable optical imaging reagent for

apoptotic cells (termed Apo-15). The functional neutrality of

Apo-15, together with the characterization of its binding to

apoptotic cells in vitro and in vivo, suggests that Apo-15 can be

used for non-invasive imaging of apoptosis in a broad range of

experimental settings, including different preclinical mouse

models.

First, we have established a solid-phase synthetic method for the

production of apopeptides (including Apo-15) as low molecular

weight (~1 kDa)

fluorogenic agents for the detection of PS, widely

regarded as an early biomarker of apoptosis. Apo-15 is shorter

than other peptides reported to bind phospholipids related to

apoptosis (e.g., PE-binding duramycin and PS-binding PSBP-6 are

19-mer and 14-mer peptides, respectively)

40

. Furthermore, Apo-15

incorporates Trp-BODIPY as an environmentally-sensitive

repor-ter for exclusive emission in proximity to cellular membranes with

enhanced brightness over other

fluorophores (Table

1

). Apo-15

can be produced cost-effectively and in large scale (>100 mgs/

batch, which would allow ~300,000 in vivo imaging assays). In

contrast, PS-binding proteins like annexins (~36 kDa) or the Ca

2+

-independent lactadherin C2 (~17 kDa) are difficult to manufacture

with site-specific chemical tags in large scale. The small size and

peptidic nature of Apo-15 might also favor better clearance and

biodistribution to tissues where proteins (e.g., annexins) cannot

gain access. For example, Apo-15 shares structural similarities (e.g.,

cyclic core, positively-charged amino acids) with recently

devel-oped blood brain barrier shuttle peptides

42,43

_{, suggesting that}

Apo-15

or slightly modified derivatives could enable optical detection of

apoptotic cells in the brain.

Second, we have demonstrated that Apo-15 binds rapidly to

aminophospholipids, with a strong preference for PS over other

phospholipids. Of note, Apo-15 binds more strongly to PS than

PE. This observation suggests that the binding to PS may be

favored by electrostatic interactions between the

positively-charged residues (i.e., arginine, lysine) of Apo-15 and the free

negatively-charged carboxylate groups in PS, which are not

pre-sent in PE. We also showed that Apo-15 binds to cells from

different species and lineages undergoing tissue culture-induced

apoptosis as well as chemical or UV-induced apoptosis (Fig.

2

).

Interestingly, although our data demonstrate that Annexin V and

Apo-15

cross-compete for binding (Fig.

3

),

fluorescence analysis

reveals that dual labeling with these two probes is not strictly

co-linear. One possibility is that there are differences in lipid

spe-cificity between Annexin V and Apo-15. Alternatively, the

assembly of Annexin V multimers on the apoptotic cell

mem-brane

31

_{may restrict accessibility to the lipids. Detection of}

membrane lipids offers key advantages (e.g., no need for

fixatives)

over optical agents visualizing other events (i.e., caspase

activa-tion, mitochondrial dysfunction or loss of membrane integrity;

Table

1

) and allows identification of apoptotic cells at early stages

of disease where therapeutic interventions may exert beneficial

effects.

Third, the divalent cation-independence of Apo-15 binding

enables in vivo detection and quantification of apoptosis in tissues

(e.g., lungs, Fig.

5

) where low and/or variable levels of free Ca

2+

could limit the performance of annexin-based reagents. Apo-15 is

also compatible with in vitro experimental conditions where the

use of annexins would be challenging, such as assays where

cal-cium ion concentrations need to be tightly controlled due to

potential activation of cells (e.g., the binding of platelets to

fibrinogen is highly dependent on the concentration of

extra-cellular free calcium)

44

_{. In addition, the enhanced}

_{fluorescence}

emission of Apo-15 upon binding allows time-lapse imaging of

apoptosis progression in situ (Supplementary Movie 1) without

the need for washing and without significant loss of signal over

time. We have exploited these characteristics to visualize the

different stages in the formation and reabsorption of apoptotic

bodies, which will provide insights into the functions that these

structures play in different diseases (Fig.

3

g and Supplementary

Movie 3).

Fourth, we have demonstrated that Apo-15 is a highly stable

peptide, even under proteolytic or oxidative conditions (Fig.

1

),

overcoming an important limitation of protein-based agents. The

low molecular weight of Apo-15 (~27-fold smaller than annexins)

may allow increased tissue penetration and biodistribution

in vivo. This feature could also enable imaging studies in

optically-transparent organisms that are well-established for

developmental studies (e.g., zebrafish (D. rerio), worms (C.

ele-gans),

flies (D. melanogaster)), given that apoptosis is crucial for

the appropriate formation of organs and structures during

embryonic development

45

_{. We have also demonstrated that the}

staining capabilities of Apo-15 are compatible with different

administration routes. Apo-15 could facilitate mechanistic and

efficacy studies of drugs in tissues undergoing repair and/or

regeneration (e.g., liver). There is experimental evidence that the

first phases of liver injury are characterized by hepatocyte

apoptosis, which is profibrogenic

46

_{. With drugs modulating}

apoptosis currently being tested for the treatment of liver injury,

the repeated in vivo administration of Apo-15—which may be

prohibitive in the case of annexins due to its cost—could

repre-sent a powerful method for longitudinal monitoring of hepatocyte

apoptosis in the liver using abdominal intravital imaging

windows

47

.

Finally, we and others have previously shown that promotion

or inhibition of apoptosis affects the progression and resolution

of inflammation

48

_{. In this work, we have demonstrated that}

Apo-15

does not accelerate or delay apoptosis, a critical requirement

for quantitative assessment of cell death in vivo. In contrast to

annexins and lactadherin C2 proteins

33,49

_{, Apo-15 does not}

inhibit the phagocytic removal of apoptotic cells by macrophages

(Fig.

4

), and therefore represents a valuable tool for imaging

assays where phagocytosis and cell clearance must remain

unaf-fected. This functional neutrality of Apo-15 may enable tracking

of the ultimate fate of apoptotic material in mice. Further studies

(we were unable to co-stain ex vivo samples with Apo-15 and

antibodies given that processing and antigen retrieval methods

results in marked loss of the Apo-15 signal) are required to

investigate whether pulses of Apo-15-labeled apoptotic cells

could be followed in vivo. Also, further studies would be required

to determine the cell types that are preferentially labeled in vivo

and the longevity of the Apo-15

fluorescence signal, particularly

when internalized apoptotic material reaches the acidic

phago-lysosome. Given experimental evidence that links dysfunctional

phagocytosis with worsening of pulmonary diseases (e.g., COPD,

cystic

fibrosis)

50

_{, Apo-15 could open avenues for imaging of}

apoptosis in other preclinical models and in clinical specimens.

Accumulation of apoptotic cells has been also reported in

atherosclerotic lesions, where defective phagocytic clearance can

contribute to the establishment of a pro-inflammatory necrotic

core and disease progression

51

. In vivo imaging of apoptosis in

these lesions, now generally done via positron emission

tomo-graphy (PET), could allow the identification of at risk vulnerable

plaques with pro-thrombotic potential

52

_{. The Trp-BODIPY in}

Apo-15

contains two

19

F

fluorine atoms which can be isotopically

exchanged with radioactive

18

_{F to prepare radiolabeled Apo-15}

analogs for multimodal imaging (i.e., PET and optical imaging)

53

_.

This approach could help to identify vulnerable plaques in vivo

and ex vivo using a single molecular agent, and also allow us to

test the efficacy of therapies promoting efferocytosis and reducing

apoptotic cell burden within atherosclerotic lesions.

In summary, our studies validate Apo-15 as a reliable

fluoro-genic peptide for the detection and quantification of apoptosis,

both in vitro in multiple cell lineages and in vivo in different

preclinical mouse models of inflammation and cancer. The

spe-cificity and non-invasiveness of Apo-15 could allow imaging of

apoptotic events that are not possible with current probes, from

tracking of apoptotic bodies with subcellular resolution to

per-forming intravital imaging using optical windows. Moreover,

Apo-15

labeling provides a simple and effective readout to

directly assess the efficacy of therapeutic interventions targeting

apoptosis in vivo.

Methods

Chemical synthesis and characterization. The synthesis and analytical char-acterization data for apopeptides is described in the Supporting Information (Supplementary Tables 1-2 and Supplementary Methods).

Cell culture and in vitro induction of apoptosis. Work with human peripheral blood leukocytes complied with all relevant ethical regulations and informed consent was obtained. The study protocol was approved by the Accredited Medical Regional Ethics Committee (AMREC, reference number 15-HV-013) at the Uni-versity of Edinburgh. Human blood leukocytes were isolated from healthy volun-teers54_{. Briefly, whole blood was anti-coagulated with sodium citrate [0.4% (w/v)}

final concentration] and centrifuged at 350 × g for 20 min. Platelet-rich plasma was removed and autologous serum was generated by recalcification with CaCl2

(20 mMfinal concentration). Erythrocytes were separated from leukocytes by sedimentation with Dextran T500 (0.6%) for 30 min and leukocyte populations were further fractionated using isotonic Percoll (GE Healthcare) 49.5%/63%/72.9% discontinuous gradients. Polymorphonuclear cells (>95% neutrophils) were har-vested from the 63%/72.9% interface. Neutrophil apoptosis was induced by in vitro culture for 18 h at 37 °C and 5% CO2in IMDM supplemented with 5% autologous

serum. Monocytes were enriched from the mononuclear cells (49.5%/63% inter-face) by negative selection using the Pan Monocyte Isolation Kit (Milentyi Biotech) according to the manufacturer’s instructions. Monocytes were cultured for 6 days in IMDM supplemented with 5% autologous serum to yield monocyte-derived macrophages (MDMs).

For work with human cell lines, BL-2 and PLB-985 cells were cultured in RPMI supplemented with 10% FBS, 100 U mL−1penicillin and 100 µg mL−1

streptomycin. Cells were plated on 8-well confocal chambers (1.25 × 105_cells/well) the day before imaging experiments. Apoptosis was induced by UV treatment (300 mJ cm−2with intervals of 6 × 50 mJ cm−2every 30 s in BL-2 cells) or by chemical treatment (staurosporine 1 µM for 3 h in BL-2 and PLB-985; staurosporine 1 µM for 6 h in primary airway epithelial cells).

Primary Small Airway Epithelial cells (HSAEC) were cultured in airway epithelial cell basal media supplemented with Bronchial Epithelial Growth Kit. Cells were plated on 8-well confocal chamber (5 × 104_{cells/well) the day before} imaging experiments. Forflow cytometry experiments, cells were detached using

0.05% trypsin-EDTA for 3 min at 37˚C and resuspended in HEPES-NaCl buffer containing 0.1% BSA and 2 mM CaCl2or 2.5 mM EDTA.

Mouse bone marrow-derived neutrophils were purified from C57Bl/6 mice using discontinuous Percoll gradients as previously described55_{. The purity of the}

neutrophil population was determined as 75–80% by analysis of cytocentrifuge preparations. Neutrophils were induced to undergo tissue culture-induced apoptosis by culturing for 18 h at 37 °C and 5% CO2in IMDM with 10% FBS.

Flow cytometry and_{fluorescence-activated cell sorting. All apopeptides were} reconstituted at 5 mM in DMSO. Peptides were used at 100 nM in HEPES-NaCl buffer containing 2 mM CaCl2, with or without 2.5 mM EDTA, and were incubated

with the cells for 10 min at r.t. beforeflow cytometry analysis, unless otherwise stated. For co-labeling experiments, AF647-Annexin V and PI were used at the indicated concentrations, with AF647-Annexin V being added to cells 10 min prior to analysis and propidium iodide being added directly before analysis. BALF analysis: 106_{cells were stained for 30 min on ice with mCD45-PE,} anti-mLy6G-PerCpCy5.5, and anti-mCD11c-Pacific Blue diluted 1:100 in HEPES-NaCl buffer. After washing, AF647-Annexin V was added at 25 nM in HEPES-NaCl buffer containing 2 mM CaCl2for 15 min prior to acquisition. Fluorescence

emission was measured on a 5 L LSRflow cytometer (BD). Excitation sources/ emissionfilters used: apopeptides (488 nm/525 nm); AF647-Annexin V (641 nm/ 670 nm); PI (560 nm/600 nm), anti-mCD45-PE (560 nm/580 nm), anti-mLy6G-PerCP-Cy5.5 (488 nm/700 nm), anti-mCD11c-Pacific Blue (405 nm/450 nm). Cells were sorted on the FACS Aria (BD), and data were analyzed with FlowJo software. Time-lapse microscopy and live-cell imaging. Time-lapse microscopy of apop-tosis in BL-2 cells was performed using a spinning disk confocal microscope (Andor) equipped with a live-cell incubation system (37 °C, 5% CO2). Live-cell

imaging of different lineages was performed on a confocal laser scanning micro-scope TCS SP8 (Leica) at 37 °C and 5% CO2. Prior to imaging, freshly isolated

neutrophils were incubated for 3 h at 37 °C in phenol red-free IMDM (Gibco) supplemented with 2.5% autologous serum and 20 µM R-roscovitine (Sell-eckchem). UV-treated BL-2 cells were incubated for 3 h at 37 °C prior to imaging. AF647-Annexin V and Hoechst 33342 were used as counterstains at the indicated concentrations. Apopeptide staining was performed by direct addition of apo-peptides at 100 nM and imaged after 10 min. Images were processed with ImageJ software.

Signal-to-noise ratios for apoptotic neutrophils labeled with FITC-Annexin V or Apo-15 were determined after image acquisition under the same laser power and gain and analysis using ImageJ. Briefly, raw images were converted into 16-bit grayscale images and averagefluorescence intensities in labeled cells and in the background were measured (≥150 points per condition). Experiments were performed in triplicate.

Liposome-based assays. Giant unilamellar vesicles (GUV) were prepared by the electroformation method56_{. Briefly, stock solutions of lipids (0.2 mM total lipid}

containing 0.3 mol% Lissamine-Rho-DOPE) were prepared in CHCl3:MeOH (2:1,

v/v) and deposited on Pt wires that were placed under vacuum for 2 h to com-pletely remove the organic solvents. 500 µL assay buffer (25 mM HEPES, 150 mM NaCl, pH 7.4) pre-warmed at 37 °C was then added to the chamber, and the Pt wires were connected to a TG330 function generator (Thurlby Thandar Instru-ments), with thefield being applied in three steps at 37 °C: (1) frequency 500 Hz, amplitude 260 mV (29 V m−1) for 6 min; (2) frequency 500 Hz, amplitude 3.1 V (320 V m−1) for 20 min; (3) frequency 500 Hz, amplitude 7.3 V (656 V m−1) for 90 min. GUVs were visualized under an inverted confocalfluorescence microscope (Nikon D-ECLIPSE C1) with the following excitation wavelengths: 488 nm for Apo-15, 561 nm for Lissamine-Rho-DOPE, and 637 nm for AF647-Annexin V. Images were collected using band-passfilters (515 ± 15 nm for Apo-15, 593 ± 20 nm for Lissamine-Rho-DOPE), and a long-passfilter >650 nm for AF647-Annexin V. For AF647-AF647-Annexin V staining, buffer containing 25 mM HEPES, 150 mM NaCl, 2 mM CaCl2, and 0.1% BSA was used. Polarizedfluorescence

emission was measured in afluorimeter plate reader PHERAstar (BMG) containing polarizers andfilters with excitation at 485 nm and emission at 520 nm. Binding of Apo-15 to phospholipids. Cardiolipidin, PG, and PA were purchased from Stratech Scientific Ltd. PS and PC were obtained from Sigma-Aldrich and PE was purchased from Cambridge Biosciences. All lipids were dissolved at 1 mg mL−1in dry EtOH and transferred into a blackflat-bottom 96-well plate to generate lipid thin layers by solvent evaporation at r.t. Apo-15 was reconstituted at 1 µM in sterile PBS and incubated at 25 °C for 40 min. Fluorescence emission was measured in a Synergy H1 BioTek spectrophotometer (λexc.: 450 nm,λexc.: 520 nm).

All experiments were performed in triplicate.

Competition binding assays between Apo-15 and AF647-Annexin V. Neu-trophils which had been induced to undergo tissue culture-induced apoptosis (cultured for 18 h at 37 °C and 5% CO2in IMDM supplemented with 5%

auto-logous serum) were pre-incubated for 10 min with different concentrations of AF647-Annexin V or Apo-15 (as indicated in Fig.3and Supplementary Fig. 12) in HEPES-NaCl buffer containing 0.1% BSA and 2 mM CaCl2. Pre-incubation was